Virus-assisted algal cell disruption for cost-effective biofuel production

ABSTRACT

An efficient virus based wet biomass lipid extraction system is disclosed herein. The system includes a microalgal biomass which is inoculated with a matched algal virus. The biomass is then incubated under conditions resulting in the algal virus infecting the algae cells and proliferating to the point that the algae cells are disrupted. This releases lipids such that they may be collected by conventional techniques, including solvent extraction. In an exemplary embodiment, the microalgal biomass comprises Chlorella sp, and the virus was Chlorovirus Paramecium bursaria chlorella virus (PBCV-1).

FIELD

The present application relates to biofuel production from wet biomass.Particularly, viral infection of microalgae is used as a natural celldisruption method for obtaining lipids used in algal biofuel production.

BACKGROUND

Biofuel production from microalgae has been advocated as one of thepotential alternative energy sources that is sustainable andenvironmentally friendly. Lipid extraction has been identified as one ofthe most cost and energy intensive steps during the production of algalbiofuel. Various disruption methods have been developed to maximize thelipid extraction, but most of them are economically infeasible toscale-up due to the requirement of extreme conditions and high costs.

This section introduces aspects that may help facilitate a betterunderstanding of the disclosure. Accordingly, these statements are to beread in this light and are not to be understood as admissions about whatis or is not prior art.

Rapid industrialization and human population growth have resulted in thedepletion of fossil fuels and rising of CO₂ emissions, which havepromoted the development of sustainable and environmental friendlybiofuels. Compared with other energy crops commonly used for biofuelproduction, microalgae have shown advantages on high growth rate, lowarable land use, and high lipid content. Lipids in microalgae, servingas the reservoir of carbon and energy in microalgal cells, are envelopedin droplets within microalgae's cytoplasm and protected by rigid algalcell walls.

In order to exploit these intracellular lipids for biofuel production,many methods have been developed for microalgal lipid extraction atlaboratory scale, but these methods are usually associated with highextraction costs. Depending on the type of microalgal biomass applied inthe extraction process, these extraction techniques can be categorizedinto dry biomass and wet biomass based extraction methods. Dryextraction is currently technologically mature but the energy intensivedrying/dehydration process has made wet extraction a competitivealternative. Wet extraction skips the energy intensive dehydration step.However, a cell disruption pretreatment is required to break rigid algalcell walls and neutralize the surface charge of algal cells to securethe lipid yield.

So far, diverse cell disruption methods based on physical, chemical, andbiological processes have been developed, but large-scale application ofthese methods is still restricted by high cost. Physical disruptionmethods such as bead-beating, high pressure homogenization (HPH),ultrasonication, microwave and electroporation have been developed tobreak cell walls physically. Chemical disruption methods depend onchemical reactions to lyse or break algal cell walls. Physicaldisruption methods demand intensive energy to support their thermal,electrical or mechanical treatments, while chemical disruption methodsare less energy intensive and easier to scale-up. However, great demandsof chemicals, need of chemical waste treatment and disposal, as well aspotential equipment corrosion remain, challenging the economicperformance of chemical disruption.

Biological disruption methods are based on the idea of enzymaticdegradation of cell walls, which can be processed under mild reactionconditions with high selectivity. Actually, various types of microalgaeeven with very resistant layers are able to be lysed by a specificmixture of enzymes at relevant energy cost. The limitation of enzymaticcell disruption, however, is the high cost of enzyme processing. Thecombination of different disruption methods such as thermal disruptionand enzymatic disruption could achieve higher lipid extractionefficiency, but the inherent constraints of the energy demand forthermal disruption and the high enzyme cost still exist.

For these reasons, innovative disruption techniques characterized by lowcost and high disruption efficiency should be developed.

SUMMARY

This disclosure provides a method of extracting lipid from a wetmicroalgal biomass. In one aspect, the method comprises inoculating thewet microalgal incubating the matched virus with the wet microalgalbiomass to initiate virus infection and replication, continuing toincubate the matched virus with the wet microalgal biomass until thevirus replication has disrupted at least 90% of the microalgae cellswithin the wet microalgal biomass, and collecting crude lipids releasedfrom the disrupted microalgae cells. In embodiments, the method used abiomass that is as much as 99% water, and the matched virus is added tothe microalgal biomass at a virus-to-algae ration of at least 1.46×10⁻⁸.The cell disruption may proceed to disrupt 99.99% or more of the algaecells in the biomass. In some preferred embodiments the method usesmicroalgal Chlorella sp in 99% water as the wet biomass and usesParamecium bursaria chlorella virus (PBCV-1).

It is an object of the present invention to provide a method forderiving lipids from algae, which has lower costs than the use ofenzymes required in biological methods.

Another object of the present invention is to obtain lipids from algaewithout requiring the continuous input of energy and chemicals common tomechanical and chemical disruption methods.

It is an advantage of the present invention that the algal viruses whichare used can be acquired and maintained with minimal costs.

Further forms, objects, features, aspects, benefits, advantages, andembodiments of the present invention will become apparent from adetailed description and drawings provided herewith.

BRIEF DESCRIPTION OF THE DRAWINGS

The patent or application file contains at least one drawing executed incolor. Copies of this patent or patent application publication withcolor drawings will be provided by the Office upon request and paymentof the necessary fee.

The above and other objects, features, and advantages of the presentinvention will become more apparent when taken in conjunction with thefollowing description and drawings wherein identical reference numeralshave been used, where possible, to designate identical features that arecommon to the figures. The attached drawings are for purposes ofillustration and are not necessarily to scale.

FIG. 1 is a picture of a plaque assay showing clear plaque forming unitsof Chlorovirus on a green layer of Chlorella sp.

FIG. 2 is a picture of gel electrophoresis to confirm the identity ofthe virus.

FIGS. 3A-3I include pictures of concentrated Chlorella cells beforeinfection (A-C), 43 hours post infection (D-F) and 54 hours postinfection (G-I). (A) Uninfected Chlorella cells after centrifugation;(B) TEM picture of uninfected cells; (C) Zoom view of an uninfected cellcorresponding to the frame in (B); (D) Centrifuged Chlorella pellet 43hours post infection; (E) TEM picture of cells 43 hours post infection;(F) Detail of one infected cell with assembled virus particles (blackarrow) and empty virus capsids (white arrow) corresponding to the framein (E); (G) Centrifuged Chlorella pellet 54 hours post infection; (H)TEM picture of lysed cells 54 hours post infection; (I) Enlarged cellsin (H), the white arrow marks the empty attached virus capsid anddegraded cell wall, and the black arrow indicates a lysed cell with ahighly deteriorated cell wall.

FIG. 4 is a graph showing infection behavior of concentrated microalgalbiomasses with different multiplicity of infection (MOI).

FIG. 5 is a graph showing viral treated and untreated efficiencies ofintracellular material release at various ultrasonication time. Errorbars represent the standard deviation of the mean.

FIG. 6 is a graph depicting FAMEs yield in pellet, supernatant and totalwith various disruption methods. Error bars represent the standarddeviation of the mean.

FIG. 7 shows the release of intracellular materials after sonicationtreatment. The white pellet (indicated by the arrow) in the left vialindicated that intracellular materials were mostly released aftersonication as the undisrupted pellet showed (green) color (middle vial).

FIG. 8 is a graph showing the absorbance spectrum of supernatant afterdifferent treatments. The peak at 675 nm was used to quantifychlorophyll released from algal cells.

DETAILED DESCRIPTION

For the purposes of promoting an understanding of the principles of thepresent disclosure, reference will now be made to the embodimentsillustrated in the drawings, and specific language will be used todescribe the same. It will nevertheless be understood that no limitationof the scope of this disclosure is thereby intended.

Host-Virus System

The present invention involves the use of a host-virus system in whichthe host cells are infected, and eventually ruptured, by a virusprovided for that purpose. A typical viral infection starts with theattachment of virus on the host cell followed by the injection of viralDNA. Then the transcription system within the host algal cell isreprogrammed by viral DNA to produce and assemble new virus particles.In particular, the present invention is directed to a host-virus systemcomprising a microalgal biomass and a matched algal virus.

Unlike other conventional disruption methods which are chemical andenergy intensive, this approach can be triggered by trace amounts ofvirus and accomplish essentially complete disruption in several daysusing normal growth conditions. With less energy input, this viraldisruption approach exhibits comparable performance to traditionaldisruption methods, which improves the extraction efficiencysignificantly from undisrupted wet biomass. Moreover, viral disruptionsignificantly reduces the mechanical strength and increases the solventpermeability of algal cells.

In general, using viral infection to disrupt algal cells does notrequire the continuous input of energy and chemicals in mechanical andchemical disruption methods. The cost of algal viruses is also lowerthan the enzymes required in biological methods. Disclosed herein aresystems and methods for the economical disruption of cells for use in amethod for biofuel production. Viral infection of microalgae is used asan economical cell disruption method of wet lipid extraction in algalbiofuel production.

In an exemplary embodiment, Chlorovirus Paramecium bursaria chlorellavirus (PBCV-1) was isolated from a natural water system and was used toinfect Chlorella sp. This system was used as it was a common and wellunderstood microalgae-algal virus system. The algal suspension hadapproximately 99% water content. An ultrasonic disruption method wasused as a reference. Viral disruption with multiplicity of infection(“MOI”) 10⁻⁸ was able to disrupt all algal cells in 6 days.

Lipid yield as well as lipid composition were determined after allmicroalgal cells were lysed. This new cell disruption method formicroalgae was determined to allow crude lipid extraction in anefficient and economic method for extracting lipids from wet algalbiomass. This also demonstrated that viral disruption can significantlyreduce the mechanical strength and increase the solvent permeability ofalgal cells.

Lipid yield with viral disruption increased more than 3 times comparedto no disruption, and was also slightly higher than the lipid yieldobtained from ultrasonic disruption alone at optimal conditions.Moreover, the quality of extracted lipids was not affected by viralinfection. Most importantly, compared to the extensive energy andchemical input required by previous disruption methods, the cost ofviral disruption is negligible. Taken together, the results confirmedthe feasibility of applying viral infection to disrupt algal cells forlipid extraction as a method for cost-efficient biofuel production. Inparticular embodiments, the system operates in the absence of otherdisruption mechanisms, or it may be supplemented with sonication orother approaches.

Microalgal Biomass

The biomass used in the present invention is a wet, microalgal biomass.The microalgae contained in the biomass may be any comprising algaecells that can be infected, and ultimately disrupted, by a matchedvirus. The wet biomass may include a wide range of percentages of water.The concentration may, for example, be at least 90% water by weight,preferably at least 95% water, and more preferably at least 99% water.

Generally, any concentration is useful, but the concentration may affectyield. Persons of ordinary skill in the art will be able to selectconcentrations suitable to make the process acceptable in terms ofdesired costs, yield, use of materials, duration and the like.

Algal Virus

The host-virus system uses a virus that is “matched” to the host algae.A matched virus is one that infects the algal cells and proliferates tothe point of disrupting (lysing) the algae cells. A matched virus is onethat meets the limitations discussed herein as to causing the celldisruption in a desire time frame.

As used herein, the term “matched virus” is not limited to a singlespecies or strain of virus for a given system. For example, differentplaques may be formed by different types of Chlorovirus as variousspecies of Chlorovirus have been detected and isolated. Thus, thematched algal virus may include a combination of two or more species orstrains which together constitute the algal virus used with a givenmicroalgal biomass. However, in order to minimize unpredictableinfection behavior caused by a mixed Chlorovirus community, theChlorovirus stock exemplified herein was established from a singleplaque and identified to be PBCV-1 with PCR.

Amount of Virus

The amount of virus added to the biomass, the “virus-to-algae” ratio,may be any amount sufficient to provide the desired extent of celldisruption within the desired amount of time. For example, in certainembodiments the matched algal virus is inoculated into the microalgalbiomass at a virus-to-algae ration of at least 1.46×10⁻⁸. As shown inFIG. 4, virus-to-algae ratios (a.k.a. multiplicity of infection (MOI))ranging from 1.46×10⁻⁸ to 1.46×10⁻² were demonstrated to be effective.Even at a very low initial MOI of 1.46×10⁻⁸, viruses still successfullyinfected algal cells with only a delay of 30 hours, because the virusesreproduced and synthesized new viruses quickly. The result of viralinfection at such a low initial concentration indicated that usingviruses for algal biomass treatment can be a robust technique forindustrial application. Continuous operation and effluent recycling canhelp maintain a high virus concentration in industrial processes.

The host-virus system we used in this work is fresh water microalgaeChlorella sp. and related virus PBCV-1. The algal biomass is treatedusing a virus effective to infect the microalgal biomass. Other algalviruses are also useful in accordance with the present method,particularly those that can infect algal species with high lipidcontent.

Incubation Parameters

The microalgal biomass and matched algal virus may be obtained inconventional fashions. The host-virus system also may be incubated usingconventional incubation equipment and operating conditions. In general,the host-virus system is operated under conditions which result in theinfection and proliferation of the algal virus to lyse the algae cells.Advantageously, these conditions may include typical incubationparameters, including temperature, pressure and other aspects known inthe art.

The time of incubation is determined based on typical factors such asthe virus-to-algae ratio of the initial inoculation, the rate ofreplication of the algal virus, the targeted end point in terms ofpercent cell disruption, etc. In accordance with the teachings herein,these parameters may be adjusted to achieve the cell disruption in thedesired time. For example, the period of incubation can be less than twoweeks, preferably less than one week, and more preferably less than 2-3days.

Cell Disruption

When the number of virus particles reaches particular level, the algalcell is lysed and virus particles are released into the aqueousenvironment. A successful infection is able to cause the lysis of algalcell in a few hours while releasing hundreds of newly assembled virusparticles that are ready for subsequent infection. Therefore, traceamounts of algal virus are able to trigger massive disruption of algalcells in ambient environment without additional input of energy andchemicals.

The incubation of the algal virus with the biomass is continued until apredetermined extent of cell disruption has occurred. In certainembodiments, for example, the process proceeds until at least 90% of thealgae cells have been disrupted, preferably at least 95% of the cells,and more preferably at least 99% of the cells. As described herein, inanother embodiment the algal virus was incubated with the biomass untilmore than 99.99% of the algae cells were disrupted.

Solvent Extraction

The purpose of the cell disruption is to facilitate recovery of lipidsfrom the algae cells, such as for use in biofuels. The released lipidsmay be collected by a variety of known techniques. The collection mayinclude using centrifugation to separate the disrupted biomass into asolid phase and a supernatant. In one approach, exemplified herein, thelipids are then recovered by solvent extraction from the supernatant. Byway of example, the lipids in the Example were captured using chloroformas the solvent. In some embodiments, the obtained crude lipids aretransesterified to the form of fatty acid methyl esters (FAMEs).

The viral disruption system was shown to provide good yields of lipids.The yield of lipids by viral disruption were compared with the yieldfrom sonication for the same batch of algal cells. Sonication is theindustry standard for disrupting algal biomass. The results showed thatviral infection was as good as sonication (no statistical differences ata significance level of 0.5). In some preferred embodiment theaforementioned method lipids yields are about 5.2±0.2% wt/wt to about4.9±0.4% wt/wt. The viral disruption technique can therefore be used toachieve good yield of lipids, while improving cost-effectiveness ascompared to existing algal processing facilities the utilize sonicationas the main extraction approach.

Example Materials and Methods

1. Microalgae Cultivation and Quantification

The host-virus system of Chlorella sp. and Chlorovirus was used todemonstrate the use of viral disruption for wet lipid extraction. Freshwater microalgal strain Chlorella sp. (ATCC 50258) was used as it is thehost of the widely spread Chlorovirus. The host-virus systems ofChlorella and Chlorella viruses have been well studied genetically andbiologically.

Stock culture was cultivated under a 14-h light, 10-h dark cycle in anenvironmental chamber at 25° C. in ATCC 847 growth medium. Light wasprovided by an IPOWER Super HPS 600 W lamp installed in theenvironmental chamber and all culture flasks were distributed at adistance of 1.1-2.1 m from the light source. ATCC medium 847 contains1.0 g Proteose Peptone, 250 mg NaNO₃, 25 mg CaCl₂), 75 mg MgSO₄, 75 mgK₂HPO₄, 175 mg KH₂PO₄, 25 mg NaCl, and a drop of 1.0% FeCl₃ solution in1.0 L distilled water. The growth of microalgae was determined by ahemocytometer (HAUSSER SCIENTIFIC) and a fluorescence microscope (NikonEclipse Ni with Plan Fluor ×40 objective). LIVE/DEAD® BacLight™Bacterial Viability Kits were used to stain live/dead cells after viralinfection. The excitation/emission wavelengths for these dyes are480/500 nm for SYTO 9 stain (live) and 490/635 nm for propidium iodide(dead). Additionally, dry algal biomass was obtained gravimetricallyafter drying at 105° C. for 12 hours in an oven until there was nosignificant change in weight (Thermo Scientific).

2. Virus Isolation and Quantification

A water sample with Chlorovirus was collected from a lake in Celery BogNature Area, West Lafayette, Ind. The collected water sample wasfiltered through a syringe filter with 0.2 μm PTFE membrane (ThermoScientific) to remove non-viral particles. Then Chloroviruses in thefiltrate were isolated by the modified plaque assay method (Van Etten,Burbank et al. 1983), which was also used to quantify virus in thisstudy. A mixture of 100 μL diluted filtrate (approximately 50 PFU per100 μL), 300 μL Chlorella cells (2×10⁸ to 4×10⁸ cells/mL), 100 μLerythromycin stock (1000 mg/L) and 5 mL of soft-agar medium was pouredonto a 1.5% agar plate. Agar medium (1.5%) and soft-agar medium (0.75%)were prepared by adding 15 g and 7.5 g agar per liter ATCC 847 mediumfor plaque assay. After a week's incubation at 25° C., viral plaquesformed on agar plate were counted. Chlorovirus from a single plaque onthe algal lawn was isolated and amplified in host culture for futureuse. To maintain algal viruses, 300 μL filtrate was added to 100 mL hostmicroalgae culture (3×10⁷ to 7×10⁷ cells/mL). After three days'incubation, new virus stock was prepared by filtering the viral richsuspension through a 0.2 μm syringe filter. The viral rich filtrateswere stored at 4° C. for future use.

The type of Chlorovirus from single plaque was identified by polymerasechain reaction (PCR). A freeze-thaw pretreatment procedure consisting ofthree cycles of heating at 95° C. for 2 mins and freezing to solid wasused to release viral DNA. Three algal virus-specific primer sets (CVMs,PBCVs, and ATCVs) from previous study (Short, Rusanova et al. 2011) wereexamined to identify the isolated Chlorovirus. Each 50 μL PCR reactioncontains 2 μL of the pretreated sample, 10 μM virus specific PCRprimers, DreamTaq DNA Polymerase, and 10× DreamTaq green buffer.Negative controls used water to substitute the sample. All PCR reactionswere performed in a Biorad 1000-Series Thermal Cycler with denaturationat 95° C. for 2 min, followed by 39 cycles of heating at 95° C. for 30s, annealing at primer-specific temperatures for 45 s, and extension at72° C. for 1 min. At the end of cycling, all PCR reactions weresubjected to a final extension step at 72° C. for 30 min. After PCR, 8μL of the reaction product were loaded into 1.5% Bio-rad Certified™agarose gel stained with SYBR safe stain and subject to electrophoresisin a Bio-Rad electrophoresis cell. The electrophoresis results wereobtained by Bio-Rad Gel Doc™ XR+ with Image Lab™ software.

3. Viral Disruption

The potential of viral infection as a disruption method was studied. Wefirst investigated the viral infection behaviors under various initialvirus loadings. Then we tested the physical strength and chemicalresistance of algal cells before and after viral infection. To obtainconcentrated algal suspensions for following experiments, algal cells atthe early stationary phase were centrifuged at 3220×G for 5 min(Eppendorf Centrifuge 5810 R and FALCON 50 mL polypropylene conicaltube) and then re-suspended in growth medium to achieve water content ofapproximately 99%. These concentrated suspensions were allocated into 1mL aliquots for further disruption treatments.

a. Dynamics of Viral Infection

The concentrated algal suspensions were inoculated with descending virusloadings and the initial virus concentrations in inoculated suspensionswere 0 (no virus inoculated), 1.66±0.04×10¹, 1.66±0.04×10³,1.66±0.04×10⁵, and 1.66±0.04×10⁷ PFU/mL, respectively. Thenconcentrations of live algal cells were measured twice a day for 6 daysuntil no live algal cells could be observed in all virus infectedsamples. Concentrations of live cells were plotted with incubation timeto obtain the infection behaviors with different initial virus loadings.

b. Electron Microscopy

To further examine the viral lead cell lysis at an ultrastructuralperspective, healthy cells and infected cells (43-hour and 54 hour postinfection) were observed under a Tecnai T12 transmission electronmicroscope (TEM). Samples were prepared by the Purdue ElectronMicroscopy Facility with modified method described by (Greiner, Frohnset al. 2009). Target algae suspensions were first centrifuged to obtainan algal pellet and were fixed with a cacodylate-buffered (pH6.8) 2%glutaraldehyde and 2% formaldehyde solution. After washing in buffer,samples were post-fixed in the same buffer with 2% OsO4. The sampleswere dehydrated in a series of graded acetone and then embedded in Embed812 Resin. At last diamond knives were used to acquire ultrathinsections. The ultrathin sections were stained with uranyl acetate andlead citrate for TEM observation.

c. Sonication

Ultrasonic treatment was employed to evaluate the effect of viralinfection on the mechanical strength of algal cells. A low frequency (20kHz) ultrasonic processor (FB-505, Fisher Scientific) equipped with a ⅛inch diameter ultrasonic horn was operated at 100 W input power asmechanical treatment. Cell breakage of healthy and virus disrupted algalsuspensions were investigated with various sonication time treatmentsranging from 0 to 420 seconds. The magnitude of cell rupture aftersonication was indicated by the absorbance measurements of releasedintracellular material in supernatant after centrifugation at 3220×G for5 min. The absorbance of released intracellular materials was measuredby a NANODROP 2000c Spectrophotometer (Thermo Scientific) at 675 nmwavelength using ATCC 847 medium as blank. All ultrasonicationtreatments were done in an ice water bath to minimize the influencecaused by temperature rise. All samples were prepared and measured intriplicate.

d. Lipid Extraction

The performance of lipid extraction by organic solvent was used toevaluate the effect of viral infection on the resistance of cells tochemical treatment. In this study, we modified the Bligh and Dyer method(Bligh and Dyer 1959, Teo, Jamaluddin et al. 2014) to extract lipidsfrom wet algal biomass. The concentrated algal suspensions were firstexposed to 3 experimental conditions: 1) no treatment, 2) viralinfection for 5 days with m.o.i. of 0.01 and 3) 300 seconds ofsonication. Then treated samples were centrifuged at 3220×G for 5 min toseparate liquid phase and solid phase. Separated upper layer supernatantand bottom pellet were both subjected to lipid extraction, respectively.Acquired pellets were re-suspended in 1 mL distilled water to match thewater volume of supernatant samples. These supernatant and pelletsamples were mixed with 2 mL chloroform and 1 mL methanol. Thechloroform layer was collected after 30 minutes of vortex, followed bycentrifugation at 3220×G for 5 min. Another two rounds of extractionwere implemented by adding another 2 mL of chloroform followed by samevortex and centrifugation processes to ensure complete extraction. Crudelipid was obtained by blow drying the lipid containing chloroform withnitrogen gas in a fume hood.

4. Lipid Transesterification and Quantification

Since the amount of algal biomass in our study was insufficient toprovide accurate gravimetric measurements, the obtained crude lipidswere transesterified to the form of acid methyl ester (FAMEs) andanalyzed through GC-MS. Crude lipids obtained from each sample weresubsequently transesterified with 2 mL methanol containing 5% sulphuricacid for 2 hours at 85° C. Hexane was used to extract FAMEs from themixture (1 mL hexane per mL mixture). The hexane layer was collectedafter 15 minutes of vortex and 5 minutes of centrifugation at 3220×G.Two more extractions by hexane were applied to guarantee the completeextraction of FAMEs. The extracted FAMEs were analyzed by a ShimadzuGC-2010 Plus equipped with a TQ8040 mass spectrometer and a ShimadzuAOC-5000 autosampler. The GC-MS was operated at a flow rate of 1.44mL·min⁻¹ using helium as the carrier gas and the source temperature ofMS was set at 200° C. Each run was started with a liquid injection of 1μL sample onto a HP-5MS (30 m×0.25 mm×0.25 μm, Angilent) column using1:10 split ratio. The oven program was held at 80° C. for 4 minutes,ramped to 235° C. at a rate of 10° C.·min⁻¹, then the rate was cut downto 5° C.·min⁻¹ until the temperature reached 300° C. At last oventemperature was held at 300° C. for 3 minutes to drive out residues incolumn. Supelco 37 component FAMEs standard mix was used to identify andquantify the FAMEs in this study. Each sample was spiked with 7.52mg·L⁻¹ Heptadecanoic Acid Methyl Ester (Ultra Scientific) as internalstandard for GC-MS analysis. It should be noted that the peaks (Fattyacids C 16:1 and C 17:1) that not included in the standard mix wereidentified by the mass spectral database from the Shimadzu GC-MS andquantified according to their nearest eluting peaks in standard mix. Thetotal lipid yield reported in this study was the sum of all detectedFAMEs contents, the lipids unable to be quantified by GC-MS were notincluded. The MS spectrum and detected FAMEs contents were reported.

5. Statistical Analysis

The total lipid yields as well as each extracted fatty acid werecompared according to the applied disruption methods using a one-wayANOVA at a significant level of α=0.05.

Results and Discussion

1. Virus Isolation and Identification.

According to FIG., the clear plaques on algal lawn were formed byChlorovirus contained in our field sample. However, different plaquesmay be formed by different types of Chlorovirus as various species ofChlorovirus have been detected and isolated so far (Van Etten andDunigan 2012). In order to minimize unpredictable infection behaviorcaused by a mixed Chlorovirus community, the Chlorovirus stock used inthis study was established from a single plaque and identified to bePBCV-1 with PCR. FIG. 2 is a picture of gel electrophoresis to confirmthe identity of the virus.

2. Viral Lysis of Algal Biomass

To the proposed viral disruption method for wet lipid extraction, wetalgal biomass with approximately 99% water content was consistently usedin this study. One of the reasons is that biomass with this watercontent could be easily obtained by some inexpensive harvest methodssuch as flocculation or sedimentation (Yoo, Park et al. 2012). On theother hand, albeit successful cell wall degradation of Chlorella sp.cells by viral infection of PBCV-1 have been observed (Meints, Lee etal. 1984, Meints, Lee et al. 1986), little information is available forthe infection behavior at high algal density that is suitable for lipidextraction. Therefore, the ultrastructure and dynamics of viralinfection at high algal density were investigated.

a. Ultrastructure of Viral Lysis

To investigate the infection at high algal density, algal suspensionswere exposed to PBCV-1 at m.o.i. of 0.01 and samples were collectedbefore infection, 43 hour post infection and 54 hour post infection forTEM observation. Successful cell infection and disruption were observedmacroscopically and microscopically (FIG. 3). Before infection,Chlorella cells were intact and able to form pellets aftercentrifugation, leaving a clear and uncolored supernatant (FIG. 3A). Asthe number of lysed cells increased, the supernatant was colored by thereleased intracellular components (FIG. 3D) and when most of the cellswere lysed the supernatant became turbid and colored (FIG. 3G).

We further investigated the effects of viral infection on Chlorella cellultrastructure using TEM. Healthy Chlorella cells with integratednucleus and undamaged cell walls were observed before the addition ofvirus (FIG. 3B,C). We started to see infected Chlorella cells 43 hoursafter the initial addition of PBCV-1 virus (FIG. 3E). The majority ofChlorella cells were still intact during this stage, but disorganizednucleus as well as virus capsids could be easily discovered in somecells (FIG. 3F). Empty (indicated by white arrow) and assembled(indicated by black arrow) virus capsids in hexagonal shape with adiameter of approximately 100 nm were identified in FIG. 3F, which wereconsistent with reported characteristics of PBCV-1 (Meints, Lee et al.1986).

Normally, cell wall lysis and release of virus particles would occurafter all viral capsids were assembled (Meints, Lee et al. 1986, VanEtten and Dunigan 2012). In the sample collected 54 hours postinfection, most Chlorella cells were lysed and free viral particles weredrifting in the solution. See FIGS. 2H and 2I. An empty virus capsid(white arrow in FIG. 3I) was found outside a segment of degraded algalcell wall with the contained viral DNA injected. The viral origin enzymewithin the virus capsid was responsible for the digesting of algal cellwall (Meints, Lee et al. 1984), which indicated that algal virus wasable to degrade algal cell walls and thus benefit lipid extraction.

We also observed a lysed algal cell (FIG. 3I black arrow) next to theempty capsid, the intracellular components were clustered together whilecell wall was hardly recognized. When zoomed out we found this was not asingle case as a majority of the lysed cells still grouped intracellularcomponents together with cell debris albeit the cell wall was highlydeteriorated. See FIG. 3H. This could also be validated by thecentrifugation result of the algal suspension. According to FIG. 3G, thelight (green) color in the supernatant and the dark (green) color in thealgae pellet indicated that only part of the chlorophyll was releasedinto the aqueous environment while most of the chlorophyll stayed withthe cell debris. A complete release of chlorophyll induced byultrasonication would show dark green color in the supernatant and whitecolor in the pellet (FIG. 7). Based on the above discussion, viralinfection showed the ability to degrade cell wall and disrupt algalcell. Furthermore, intracellular materials such as lipids andchlorophyll tend to stay with lysed cells instead of releasing into theaqueous environment after viral disruption.

b. Infection Dynamics for Various Multiplicities

Viral infection works in a similar way to an autocatalysis reaction, soseveral cycles of replication are necessary to accumulate a sufficientamount of virus to infect all algal cells. It is obvious that increasingMOI can reduce the cycles of replication required in the disruptionprocess, and thus reduce the processing time. However, increasing theMOI would add cost to the preparation of virus that is used for initialinoculation.

In order to balance the cost and processing time, the characteristics ofviral life cycle such as replication time and virus burst size should beconsidered. For example, fast viral replication and large burst size canreduce the time used for virus accumulation. However, thesecharacteristics are affected by inoculation conditions such as MOI andgrowth stage of host cells. Increasing MOI would cause the decreasing ofburst size, while the growth stage of host cells would affect the speedof replication (Van Etten, Burbank et al. 1983). Normally, PBCV-1 canfinish one replication in 6-8 hours and has a burst size of severalhundred PFU (Van Etten and Dunigan 2012), but these characteristics aresubject to change during the disruption process due to the increasingconcentration of virus and high concentration of algae. Therefore, theinfection behaviors with the proposed algal biomass content wereinvestigated under various MOI.

The concentrated algal suspensions were exposed to PBCV-1 loadings from1.66±0.04×10¹ PFU/mL to 1.66±0.04×10⁷ PFU/mL. As illustrated in FIG. 4,the concentration of live algal cell in the control sample (with novirus addition) remained stable throughout the experiment, whilecomplete disruption (no live cell) was achieved in all viral infectedsamples. Viral disruption could be triggered regardless of MOI but thedisruption time was observed to be inversely proportional to the appliedMOI. Under the infection conditions applied in this study, the time usedfor complete disruption ranged from 105 hours with highest MOI (1.46×10⁻²) to 142 hours with lowest m.o.i. (1.46×10⁻⁸). The increase of MOIby 6 orders of magnitude only saved 26% time used on disruption, whichindicated low MOI might be a good choice for viral disruption. In realapplication of viral disruption, the virus rich lysate after disruptioncould be recycled and fed back to disruption tank, which can reduce thedisruption time and save cost on virus preparation.

3. Viral Disruption for Lipid Extraction.

Based on the above discussion, although most of the oil is trapped inlysed cells after cell disruption, the highly deteriorated cell wall mayno longer be a tough barrier for lipid extraction. To further evaluatethe potential of using viral infection as a cell disruption method forlipid extraction from algal biomass, the mechanical strength and solventpermeability of algal cells after viral disruption were investigated.

a. Effect of Viral Disruption on Cell's Mechanical Strength

Since algal cells with reduced mechanical strength are more vulnerableto mechanical power, less energy is required for mechanical treatmentsto break cell walls and release intracellular lipids. The reduction inenergy demand is beneficial to the overall economics of the lipidextraction process. In this study, sonication was employed to evaluatethe effect of viral disruption on mechanical strength of algal cells.Sonication is able to break algal cells and cause the release ofintracellular materials that tend to stay in the supernatant rather thanprecipitate in the pellet after centrifugation (illustrated in FIG. 7).In addition, the release of intracellular materials has been used toevaluate the level of cell breakage for C. reinhardtii (Gerde,Montalbo-Lomboy et al. 2012). Therefore, the amount of releasedintracellular materials was used as a means to quantify the mechanicalstrength of algal cells under various levels of sonication. Thecharacteristic peak of released intracellular materials was located at675 nm (the absorbance spectrum is presented in FIG. 8). Untreated andvirus disrupted samples were subject to 420 and 150 seconds ofultrasonic treatments, respectively, until the absorbance measurementsof supernatants were stabilized. The average of the last threeabsorbance measurements were used as the maximum absorbance when allintracellular materials were released. The efficiency of cell breakageat each sonication energy level was then calculated by dividing theabsorbance reading by the maximum absorbance.

The efficiency of cell breakage was plotted with sonication energy foruntreated and virus disrupted samples (FIG. 5). For both untreated andvirus disrupted samples, the efficiency of cell breakage surged at lowenergy levels, then the increase of efficiency slowed down as sonicationenergy increased, and finally the increase ceased when all cells weredisrupted. Although the cell breakage patterns of untreated and virusdisrupted samples were similar, the virus disrupted cells showed greatreduction on their mechanical strength. To achieve 80% of cell breakage,the sonication energy for virus disrupted samples was 1.5 kJ/mL, whichwas 8.3% of the sonication energy 18 kJ/mL required for untreatedsamples. As for a complete cell breakage, the necessary sonicationenergy for virus disrupted samples was 4.5 kJ/mL, which was 15% of the30 kJ/mL energy needed for untreated samples. According to theseresults, sonication appeared to be more efficient to releaseintracellular materials in virus disrupted suspensions, especially atlow energy levels. As observed in FIGS. 3H and 2I, the virus disruptedalgal cells were highly deteriorated and the crumbled cell wall couldbarely hold the intracellular materials in the cell envelope. Therefore,instead of consuming additional energy to break rigid cell wall, theintracellular materials in virus disrupted cells just need to beagitated out of the damaged cell envelope by sonication. Both theultrastructure of virus disrupted cells as well as the results fromsonication test indicated that viral disruption could significantlyreduce the mechanical strength of algae cells.

b. Lipid Extraction with Organic Solvent

Lipids extraction via organic solvents was used to evaluate theperformance of viral disruption, while no disruption and ultrasonicdisruption were applied as the reference to the lower and upper limitsof the performance. Lipids from the biomass pellet and the supernatantwere analyzed separately to provide information about the distributionof lipids when different disruption methods were applied. As describedpreviously, only detected FAMEs contents were used to calculate lipidyield. The highest lipid yields, 5.2±0.2% wt/wt and 4.9±0.4% wt/wt, wereobtained by using both the viral disruption method and the ultrasonicdisruption method (FIG. 6) as there was no statistically significantdifference between the lipid yields from these two methods. However, thelipid yield with no treatment was 1.5±0.3% wt/wt, which was only about30% of the lipid yield when disruption methods were applied. The lowlipid yield from undisrupted algal cells by applying direct solventextraction was due to the surface charges of healthy algal cells, whichkept them within water phase and prevented sufficient contact with thesolvent (Kim, Yoo et al. 2013). On the contrary, algal cells subjectedto ultrasonic disruption were ruptured into small pieces of cell debris,which eliminated the barrier between intracellular materials and organicsolvent. Similarly, the virus disrupted cells that lost their surfacecharges and cell integrity could no longer impede the contact betweenintracellular lipids and solvent.

Viral disruption has demonstrated the ability to perform comparable celldisruption as sonication, however, lipids tend to be held within lysedcells after viral disruption rather than dispersed into the solutionafter ultrasonic disruption. According to the results reported in (FIG.6). 100% and 95% of the lipids extracted with no disruption and viraldisruption were from the biomass pellet, while over 97% of the lipidyield with ultrasonic disruption came from supernatant. In samples withno disruption, lipids were enveloped in intact cells thus they wouldundoubtedly precipitate to the bottom with the biomass. The ultrasonicprocess is capable of breaking cells into small pieces that are unableto be precipitated by centrifugation, and the cell debris can trapreleased intracellular materials such as lipids (Wang and Wang 2011,Gerde, Montalbo-Lomboy et al. 2012), which explained the high lipidcontent observed in the supernatant of ultrasonic disrupted samples. Forvirus disrupted samples, the distribution of lipid showed similar trendto undisrupted samples. Although the virus disrupted cell walls werehighly damaged, lipids tended to stay in lysed cells rather than bereleased into the solution, which was consistent with the analysis withTEM as described before.

c. Effect of Viral Disruption on Composition of Fatty Acids

The viral infection process can reprogram the metabolic pathway of thehost algae, which may affect the composition of extracted lipids. Forexample, it has been reported that a marine algal virus EhV is able toreprogram the metabolic pathway of their host E. huxleyi to synthesizemore triglycerides (TAGs) after viral infection (Rosenwasser, Mausz etal. 2014, Malitsky, Ziv et al. 2016). Furthermore, algal virusesthemselves are a possible source of lipids. PBCV-1 has been reported tocontain a lipid membrane and another virus EhV that can infect E.huxleyi was reported to possess high content of TAGs (Van Etten andDunigan 2012, Malitsky, Ziv et al. 2016).

TABLE 1 Fatty acids composition of extracted lipids with differentdisruption treatments Fatty FAME yield (10⁻³ mg FAME /mg dry biomass)acids No Treatment Sonication Viral disruption C 15:0 1.21 ± 0.086  2.71± 0.023  2.77 ± 0.067 C 16:0 2.33 ± 0.649  9.24 ± 1.359  9.27 ± 0.458 C16:1 4.11 ± 0.057 11.33 ± 0.317 11.98 ± 0.380 C 17:0 1.80 ± 0.658  5.99± 0.647  7.65 ± 1.207 C 17:1 1.30 ± 0.011  4.17 ± 0.099  4.72 ± 0.359 C18:2 1.30 ± 0.685  4.97 ± 1.014  4.79 ± 0.202 C 18:3 2.58 ± 0.946 10.64± 0.976 10.84 ± 0.596

The FAMEs contents obtained from different disruption methods wereanalyzed statistically and viral disruption did not show significantinfluence on lipid contents. The FAMEs profiles for each disruptionmethod and the detailed values are listed in Table 1.

The total yields obtained from both the supernatant and the pellet foreach type of FAME were used to generate the FAMEs profile. Thestatistical analysis for each fatty acid was summarized in Table 2.According to the results, the yields of all types of fatty acids withoutdisruption were much lower than the yields with disruption, which couldbe attributed to the low extraction efficiency caused by the intact cellwall. On the other hand, similar fatty acids profiles were obtained fromboth ultrasonic and viral disruption methods, which indicated that viraldisruption was able to perform the disruption as efficiently asultrasonic disruption without affecting the quality of extracted lipids.However, it is worth noting that the yields of fatty acids C 16:0, C17:0, and C 17:1 obtained with viral disruption were higher than thoseobtained with ultrasonic disruption at an almost significant level.

The p-values obtained from an ANOVA test for these three fatty acidswere 0.085, 0.062, and 0.104, respectively, which were close to thesignificance level of 0.05, showing that viral infection may have theability to affect the quality of lipids in a way that is favorable tobiofuel production.

TABLE 2 Statistical analysis results of ANOVA analysis of individualfatty acids p-value of ANOVA tests No No treatment treatment SonicationFatty and and viral and viral acids sonication disruption disruption C15:0 0.002 0.002 0.217 C 16:0 <0.001 <0.001 0.085 C 16:1 0.001 <0.0010.973 C 17:0 <0.001 <0.001 0.062 C 17:1 0.001 0.002 0.104 C 18:2 0.005<0.001 0.773 C 18:3 <0.001 <0.001 0.778

Nonetheless, the relatively low lipid yield with ultrasonic treatmentmay be due to the sonication induced lipid oxidation (Gerde,Montalbo-Lomboy et al. 2012). Therefore, the lipid content was notaffected by viral disruption in this study, however, more in-depthinvestigations should be conducted in the future to explore the impactof virus on the lipid metabolic pathway of host algae.

4. Economical Aspects In addition to the comparable disruptionperformance, using viral lysis to disrupt wet algal biomass for lipidextraction has shown significant economic potential over other existingdisruption methods reported in literature. The infection and lysisprocesses of algal biomass can be operated under normal temperature andpressure in wet biomass. The isolated virus massively proliferates in ashort time to meet the demand of treatment. Part of the virus richlysate after disruption can be recycled and fed to fresh wet biomass tosave the time and cost in virus preparation. Therefore, viral disruptiondoes not require the extensive energy and chemical input applied inphysical and chemical disruption methods. Additionally, algal virusescan be acquired and maintained with minimal costs as compared to thehigh costs of enzymes that are available for cell wall degradation.Consequently, the moderate operation conditions and low cost of reagentshave made viral disruption a competitive choice in large scale lipidextraction systems.

Those skilled in the art will recognize that numerous modifications canbe made to the specific implementations described above. Theimplementations should not be limited to the particular limitationsdescribed. Other implementations may be possible.

1. A method of extracting lipids from a wet microalgal biomass,comprising: inoculating the wet microalgal biomass with a matched virus;incubating the matched virus with the wet microalgal biomass to initiatevirus infection and replication; continuing to incubate the matchedvirus with the wet microalgal biomass until the virus replication hasdisrupted at least 90% of the microalgae cells within the wet microalgalbiomass; and collecting crude lipids released from the disruptedmicroalgae cells.
 2. The method of claim 1 in which said inoculatingcomprises adding the matched virus to the microalgal biomass at avirus-to-algae ratio of at least 1.46×10⁻⁸.
 3. The method of claim 2 inwhich said inoculating comprises adding the matched virus to themicroalgal biomass at a virus-to-algae ratio of 1.46×10⁻⁸ to 1.46×10⁻².4. The method of claim 3 and which includes continuing to incubate thematched virus with the wet microalgal biomass until at least 99% of themicroalgae cells have been disrupted.
 5. The method of claim 4 in whichsaid continuing to incubate is performed under conditions and for a timeeffective to disrupt at least 99.99% of the algae cells.
 6. The methodof claim 5 in which the wet microalgal biomass comprises at least 99% byweight water.
 7. The method of claim 1 in which said collectingcomprises solvent extracting the crude lipids.
 8. The method of claim 1and which, following the continuing step and preceding the collectingstep, includes using centrifugation to separate the biomass into a solidphase and a liquid supernatant.
 9. The method of claim 8 in which saidcollecting comprises solvent extracting the crude lipids from thesupernatant.
 10. The method of claim 1 in which the wet microalgalbiomass is microalgal Chlorella sp in 99% water.
 11. The method of claim10 in which the matched virus is Paramecium bursaria chlorella virus(PBCV-1).
 12. The method of claim 11 in which said collecting comprisessolvent extracting the crude lipids.
 13. The method of claim 12 in whichsaid solvent extracting uses chloroform as the solvent.
 14. The methodof claim 13, and further including transesterification of the obtainedcrude lipids to the form of acid methyl esters (FAMEs).
 15. A method ofextracting lipids from a wet microalgal biomass, comprising: inoculatingthe wet microalgal biomass with a matched virus at a virus-to-algaeratio of at least 1.46×10⁻⁸, the wet microalgal biomass comprising atleast 99% water; incubating the matched virus with the wet microalgalbiomass to initiate virus infection and replication; continuing toincubate the matched virus with the wet microalgal biomass until thevirus replication has disrupted at least 99% of the microalgae cellswithin the wet microalgal biomass; and solvent extracting crude lipidsreleased from the disrupted microalgae cells.
 16. The method of claim 15in which, following the continuing step and preceding the solventextracting step, includes using centrifugation to form a solid phase anda liquid supernatant, said solvent extracting being performed on thesupernatant.
 17. The method of claim 16 in which the wet microalgalbiomass is microalgal Chlorella sp in 99% water.
 18. The method of claim17 in which the matched virus is Paramecium bursaria chlorella virus(PBCV-1).
 19. The method of claim 18 in which said solvent extractinguses chloroform as the solvent.
 20. The method of claim 19, and furtherincluding transesterification the obtained crude lipids to the form ofacid methyl esters (FAMEs).